Repurposing Loperamide as an Anti-Infection Drug for the Treatment of Intracellular Bacterial Pathogens

Hongtao Liu; Siqi Li; Le Deng; Zhenxu Shi; Chenxiao Jiang; Jingyan Shu; Yuan Liu; Xuming Deng; Jianfeng Wang; Zhimin Guo; Jiazhang Qiu

doi:10.1016/j.eng.2024.01.011

Engineering ›› 2024, Vol. 39 ›› Issue (8) :193 -206. DOI: 10.1016/j.eng.2024.01.011

Research

Article

Repurposing Loperamide as an Anti-Infection Drug for the Treatment of Intracellular Bacterial Pathogens

Hongtao Liu ^a^,^b^,^#
, Siqi Li ^a^,^b^,^#
, Le Deng ^a^,^b
, Zhenxu Shi ^a^,^b
, Chenxiao Jiang ^a^,^b
, Jingyan Shu ^a^,^b
, Yuan Liu ^c
, Xuming Deng ^a^,^b
, Jianfeng Wang ^a^,^b
, Zhimin Guo ^a^,^b
, Jiazhang Qiu ^a^,^b^,^*

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History +

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Abstract

Infections caused by intracellular bacterial pathogens are difficult to treat since most antibiotics have low cell permeability and undergo rapid degradation within cells. The rapid development and dissemination of antimicrobial-resistant strains have exacerbated this dilemma. With the increasing knowledge of host-pathogen interactions, especially bacterial strategies for survival and proliferation within host cells, host-directed therapy (HDT) has attracted increased interest and has emerged as a promising anti-infection method for treating intracellular infection. Herein, we applied a cell-based screening approach to a US Food and Drug Administration (FDA)-approved drug library to identify compounds that can inhibit the intracellular replication of Salmonella Typhimurium (S. Typhimurium). This screening allowed us to identify the antidiarrheal agent loperamide (LPD) as a potent inhibitor of $S$ . Typhimurium intracellular proliferation. LPD treatment of infected cells markedly promoted the host autophagic response and lysosomal activity. A mechanistic study revealed that the increase in host autophagy and elimination of intracellular bacteria were dependent on the high expression of glycoprotein nonmetastatic melanoma protein B (GPNMB) induced by LPD. In addition, LPD treatment effectively protected against $S$ . Typhimurium infection in Galleria mellonella and mouse models. Thus, our study suggested that LPD may be useful for the treatment of diseases caused by intracellular bacterial pathogens. Moreover, LPD may serve as a promising lead compound for the development of anti-infection drugs based on the HDT strategy.

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Keywords

Intracellular bacteria / US Food and Drug Administration (FDA)-approved drugs / Drug repurposing / Loperamide / Autophagy / Glycoprotein nonmetastatic melanoma / protein B

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Hongtao Liu, Siqi Li, Le Deng, Zhenxu Shi, Chenxiao Jiang, Jingyan Shu, Yuan Liu, Xuming Deng, Jianfeng Wang, Zhimin Guo, Jiazhang Qiu. Repurposing Loperamide as an Anti-Infection Drug for the Treatment of Intracellular Bacterial Pathogens. Engineering, 2024, 39(8): 193-206 DOI:10.1016/j.eng.2024.01.011

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1. Introduction

Infections caused by intracellular bacterial pathogens such as Salmonella enterica, Mycobacterium tuberculosis (M. tuberculosis), Shigella flexneri (S. flexneri), and Listeria monocytogenes (L. monocy-togenes) result in millions of deaths annually worldwide and pose major public health concerns [1]. For example, approximately ten million people contract tuberculosis (TB) caused by M. tuberculosis, leading to an estimated 1.5 million deaths per year [2], and Salmonella enterica causes approximately 1.3 billion annual infections, leading to salmonellosis and posing a global health problem [3]. Traditional antimicrobial treatments have greatly contributed to patient outcomes and reduced the mortality of bacterial infections. However, the increase in bacterial resistance has markedly decreased the therapeutic efficacy of most conventional antibiotics. In particular, the emergence and rapid transmission of the mobilized colistin resistance (MCR) gene [4], New Delhi metallo-

β

-lactamase (NDM-1) gene [5], and tigecycline resistance gene tet

X 4 / 5

[6,7] are now threatening the clinical usage of the last resort antibiotics colistin, carbapenem, and tigecycline, respectively. Moreover, the elimination of bacteria residing intracellularly requires efficient uptake of antibiotics into host cells, which adds an additional level of complexity for the treatment of related infections [8]. It has been estimated that ten million people will die every year due to antimicrobial resistance by 2050 [9]. Therefore, novel and creative strategies or drugs are urgently needed to control bacterial diseases.

The intracellular environment of host cells is hostile to bacteria. However, many intracellular pathogens have evolved distinct strategies to compromise host innate antimicrobial mechanisms, thus allowing bacterial survival and successful replication within cells [10]. For instance, intracellular Salmonella utilizes the type III secretion system (T3SS) to deliver the effector protein SopF into the host cytosol; this protein functions as a general xenophagy inhibitor by targeting the V-type ATPase (V-ATPase)-autophagy-related protein 16-1 (ATG16L1) axis via adenosine diphosphate (ADP)-ribosylation, thus preventing bacteria from being eliminated by the xenophagy pathway [11]. The S. flexneri T3SS substrate invasion plasmid antigen H9.8 (IpaH9.8) targets human guanylate binding protein-1 (hGBP1) for proteasomal degradation by the 48th lysine (K48)-linked polyubiquitination, thereby conferring antibacterial defense and promoting

S

. flexneri replication in the host cytosol [12]. These advances in the understanding of host-pathogen interactions have facilitated the development of host-directed therapy (HDT) to control infections attributed to intracellular pathogens [13]. Unlike antibiotics, which aim to directly kill or inhibit bacteria by targeting the bacterial elements required for growth, HDT drugs function to eradicate intracellular pathogens by direct modulation of host cell functions [10,13].

The de novo discovery and development of novel anti-infection drugs is costly and time-consuming. In contrast, drug repurposing using US Food and Drug Administration (FDA)-approved drugs or compounds that failed in clinical trials is a reliable and efficient approach to accelerate drug development [14]. In this study, we initiated drug screening to search for inhibitors of the infection control (ST)-related intracellular replication from an FDA-approved compound library. These attempts have allowed us to identify loperamide (LPD), a common antidiarrheal agent, as an HDT candidate for treating ST infection. We found that LPD treatment could significantly augment the host autophagic response as well as lysosomal activity. In addition, LPD-treated cells exhibited elevated transcription and expression of glycoprotein non-metastatic melanoma protein B (GPNMB), which is required for the LPD-stimulated autophagic response and elimination of intracellular bacteria. Finally, the administration of LPD to animals infected with ST had marked protective effects on the basis of the observed mortality, organ bacterial load, histopathological data, and cytokine levels. Taken together, our findings support the repurposing of LPD as a drug for the treatment of intracellular pathogen-caused diseases.

2. Materials and methods

2.1. Bacterial strains, cell lines, culture methods, and chemicals

The bacterial strains used in the present study are listed in Table S1 in Appendix A. Salmonella strains were cultured at 37 °C in Luria-Bertani (LB) broth supplemented with suitable antibiotics. To induce the expression of the Salmonella Pathogenicity Island 1 (SPI-1) gene, the cultures were supplemented with

0.3 m o l ⋅ L - 1

NaCl. Escherichia coli (E. coli), S. flexneri, Acinetobacter baumannii (A. baumannii), L. monocytogenes, and Staphylococcus aureus (S. aureus) strains were grown at

37 ∘ C

in brain heart infusion (BHI) broth supplemented with suitable antibiotics.

Human cervical cancer cells (HeLa), mammary alveolar cells-large T antigen cells (MAC-T), and colorectal adenocarcinoma cells (Caco-2) were grown in Dulbecco’s modified Eagle’s medium (DMEM; Sigma-Aldrich, USA) supplemented with

10 %

fetal bovine serum (FBS), while mouse leukemic monocyte/macrophage cell line (RAW264.7) were maintained in Roswell Park Memorial Institute (RPMI) 1640 (Sigma-Aldrich) supplemented with

10 %

FBS and

1 %

penicilin/streptomycin. All cells were cultured at

37 ​ ∘ C

in an incubator containing

5 % C O 2

The FDA-approved drug library (Cat#L1021; APExBIO, USA) consists of 2320 chemical compounds.

2.2. Minimum inhibitory concentration (MIC) determination and in vitro growth curve

The MIC of LPD against the tested bacterial strains was evaluated following the Clinical Laboratory Standards Institute (CLSI) broth microdilution protocol with some modifications. Resazurin was added to the wells, and the optical density (OD)

570 n m

(OD570nm) was determined by a modular multimode microplate reader (Synergy H1; BioTek, USA).

For the in vitro growth curve assay, overnight bacterial cultures were diluted in fresh LB to obtain an OD600nm of 0.1. Aliquots of the diluted culture were supplemented with increasing concentrations of LPD and grown at

37 ∘ C

with constant rotation at 220

r ⋅ m i n - 1

. The bacteria were cultured for

5 h

, after which the OD600nm was measured at 30 min intervals.

2.3. Cell-based infection screening assay

Cells

1 × 105

were seeded in 24-well plates. ST was grown in high-salt

0.3 m o l ⋅ L - 1 N a C l L B

medium to the logarithmic phase

O D 600 n m = 1

and used to infect seeded cells at a multiplicity of infection (MOI) of 20 for

1 h

. Then, the infected samples were treated with

100 μ g ⋅ m L - 1

gentamycin for

1 h

to eliminate and remove extracellular bacteria. After the cells were washed with warm phosphate buffered saline (PBS) three times, fresh culture medium containing

40 μ g ⋅ m L - 1

individual FDA-approved drugs was added, and the cells were further cultured at

37 ∘ C

for

4 h

. Infected cells were then lysed with

0.2 %

saponin, and the lysates were spread on LB plates to determine the number of colony-forming units (CFUs).

E. coli, S. flexneri, A. baumannii, L. monocytogenes, and S. aureus were grown in BHI broth to the logarithmic growth phase

O D 600 n m = 1

and used to infect RAW264.7 cells at an MOI of 20 for

1 h

. Then, the intracellular replication of these bacteria in the presence of LPD was determined according to the procedure used for ST.

2.4. $β$ -lactamase (TEM) reporter assay

Effector translocation of the ST-T3SS-1 and T3SS-2 strains was determined via the use of a TEM reporter assay as previously described [15,16]. SipA-TEM or SopD2-TEM was subsequently transformed into wild-type S. Typhimurium strain SL1344. As negative controls, the plasmids were also introduced into T3SS-1- deficient

Δ inv A

and T3SS-2-deficient

Δ ssa N

cells. Overnight ST cultures were diluted 1:30 in fresh LB medium containing

0.3 m o l ⋅ L - 1 N a C l

. Following the addition of increasing concentrations of LPD, the bacteria were further grown for

4 h

with constant rotation at

220 r ⋅ m i n - 1

. HeLa cells were plated in 96-well plates at

1.2 × 104

cells per well and infected with ST at an MOI of 20 for

2 h

. The cells were washed twice with Hank’s balanced salt solution (HBSS) to remove extracellular bacteria. Then, a

6 ×

CCF4/AM reaction mixture (K1095; Thermo Fisher, USA) was added to each of the wells. After incubation at room temperature (RT) in the dark for 1

h

, the translocation of the effector proteins was visualized under a fluorescence microscope (Olympus IX83, USA).

2.5. Trichloroacetic acid (TCA) precipitation

Secretion of ST-T3SS-1 into the culture supernatants was determined by TCA precipitation as described previously [17]. Briefly, wild-type SL1344 was cultured in high-salt LB medium supplemented with various concentrations of LPD for

5 h

. Two milliliters of the bacterial culture was centrifuged at

12000 g

for

5 m i n

. The bacterial pellets were resuspended in

100 μ L

1 ×

sodium dodecyl sulfate (SDS) loading buffer. The supernatants were mixed with TCA to a final concentration of

10 %

and incubated overnight at

4 ​ ∘ C

. After centrifugation at

12000 g

for

20 m i n

, the precipitates were resolved in

20 μ L

1 ×

SDS loading buffer. After separation by SDS-polyacrylamide gel electrophoresis (PAGE), the proteins were analyzed via Coomassie brilliant blue (CBB) staining or western blotting using specific antibodies.

2.6. RNA sequencing and quantitative real-time polymerase chain reaction (qRT-PCR)

RAW264.7 cells were either left uninfected or infected with wild-type SL1344 at an MOI of 20 for

1 h

. After gentamycin treatment as described earlier,

16 μ g ⋅ m L - 1

LPD was added to the infected samples and incubated for

4 h

. Then, the cells were collected and subjected to total RNA isolation using TRIzol

​ T M

reagent (Cat#15596018; Thermo Fisher) according to the manufacturer’s procedure. Transcriptome analysis was performed by

2 × 150

base pair (bp) paired-end sequencing (PE150) on an Illumina NovaSeq

​ T M

6000 (LC-Bio Technology Co., Ltd., China) following the vendor’s recommended protocol. The threshold for selecting differentially expressed genes (DEGs) was a fold change (FC)

≥ 2

< 0.5

(the absolute value of

l o g 2 F C ≥ 1

). A false discovery rate (FDR)

< 0.05

was used as the criterion for screening DEGs. Bioinformatic analysis was performed using the OmicStudio tool.

Total RNA was reverse transcribed into complementary DNA (cDNA) using a RevertAid RT reverse transcription kit (Cat#K1691; Thermo Fisher) and used as a template for qRT-PCR analysis. The sequences of the primers used are listed in Table S2 in Appendix A, and qRT-PCR was carried out using synergetic binding reagent (SYBR) Green fluorescent dye (Cat#KTSM1401; AlpaLife, China) on an Applied Biosystems Real-Time PCR instrument (QuantStudio 1; Thermo Fisher). The mouse glyceraldehyde 3-phosphate dehydrogenase (GAPDH) was used as an internal control to normalize gene expression. Relative gene expression was determined according to the 2-ΔΔcycle-threshold (CT) method.

2.7. Knockdown of glycoprotein nonmetastatic melanoma B gene (gpnmb) by RNA interference

RAW264.7 cells were seeded into six-well plates

2 × 105

cells per well) and grown for

18 h

. Then, the cells were transfected with

50 μ m o l ⋅ L - 1

gpnmb small interfering RNA (siRNA; Table S3 in Appendix A) for

48 h

using Advanced DNA RNA Transfection Reagent (AD600075; Zeta Life, USA). The efficiency of gpnmb knockdown was evaluated by western blot analysis using an anti-GPNMB antibody.

2.8. Western blot and immunostaining analysis

After ST-infected RAW264.7 cells were treated with or without

16 μ g ⋅ m L - 1

LPD for

4 h

, the cells were collected and lysed with octylphenoxypolyethoxyethanol (NP-40) lysis buffer (150 mmol·L

​ - 1

NaCl,1%NP-40,50 mmol·L

​ - 1

tris-HCl, pH = 8.0). Equal amounts of protein were separated by SDS-PAGE and then transferred to nitrocellulose membranes. After blocking with 5% skim milk in tris buffered saline with Tween-20 (TBST) buffer for

1 h

, the membranes were probed with the appropriate primary antibodies for

1 h

at RT. The antibodies used (and their dilutions) were as follows: rabbit anti-SipA (1:1000), rabbit anti-isocitrate dehydrogenase (ICDH) (1:5000, Cat#abs2090; Sigma-Aldrich), rabbit anti-p62 (the protein encoded by sqstm1 gene) (1:3000, Cat#ab109012; Abcam, UK), rabbit anti-microtubule-associated protein 1 light chain 3 beta (LC3B) (SAB5701328, 1:3000; Sigma-Aldrich), rabbit anti-mammalian target of rapamycin (mTOR; 1:1000, Cat#2983; CST, USA), rabbit anti-phosphorylated mTOR (1:1000, Cat#2976; CST), rabbit anti-GPNMB (1:3000, Cat#ab188222; Abcam), and rabbit anti-tubulin (1:5000, Cat#ab210797; Abcam). The membranes were subsequently washed three times with TBST and incubated with appropriate secondary antibodies (ab175775 and ab175781; Abcam). The membranes were scanned by an Odyssey CLx Imaging System (LI-COR, USA). The protein abundance was quantified by ImageJ.

For immunostaining, cells were fixed at RT for

30 m i n

with

4 %

paraformaldehyde and permeabilized with

0.02 % v / v

Triton X- 100 for 5 min. After the cells were blocked with

4 %

goat serum for

30 m i n

37 ∘ C

, the samples were stained with the indicated primary antibodies for

1 h

at RT. The antibodies used (and their dilutions) were as follows: mouse anti-Salmonella antibody (ab69238, 1:1000; Abcam), rabbit anti-lysosomal associated membrane protein 1 (LAMP-1) (1:1000, Cat#ab24170; Abcam) and rabbit anti-LC3B (1:1000, Cat#SAB5701328; Sigma-Aldrich). Then, the cells were stained with Alexa Fluor 594- or Alexa Fluor 488- conjugated secondary antibodies (Thermo Fisher). The nuclei were visualized by Hoechst 33342 staining. Immunofluorescence signals were observed under an Olympus IX83 microscope.

2.9. Experimental treatment with LPD in ST-infected models

The animal study was reviewed and approved by the Institutional Animal Care and Use Committee of Jilin University (permit number: SY202306059).

(1) Galleria mellonella (G. mellonella) infection model. The larvae of G. mellonella were purchased from Tianjin Huiyude Biotechnology Co., Ltd. (China) and were randomly assigned to three groups (

n = 10

per group). The larvae were injected with

10 μ L

of bacterial suspension containing

1 × 105

CFUs of SL1344 in the right posterior gastropod using a syringe. Subsequently, the larvae in the treatment group were injected with

150 μ g ⋅ k g - 1

LPD dissolved in

10 μ L

of solvent

(10 %

dimethyl sulfoxide,

40 %

PEG400, 45% saline, and 5% Tween-80). Larvae in the infection control group were injected with an equal volume of the solvents. Survival of

G

. mellonella was monitored for

12 h

(2) Mouse infection model. BALB/c mice (female, 6-8 weeks old, 18-20 g) were purchased from Liaoning Changsheng Biotechnology Co. (China). Mice were randomly divided into the SL1344 infection group, LPD treatment group, and PBS group

(n = 10

per group for survival rate and

n = 5

per group for determination of organ bacterial burden). All mice were fed water containing streptomycin

5 g ⋅ L - 1

for

3 d

. On the fourth day, the infection group and the LPD treatment group were orally infected with SL1344 suspensions

1 × 107

CFUs for survival rate determination and

5 × 106

CFUs for determination of organ bacterial colonization). Mice in the treatment group were intraperitoneally administered

5 m g ⋅ k g - 1

LPD at

12 h

intervals. Mice were dissected on the eighth day, and tissue samples from the spleens and livers were homogenized in phosphate-buffered saline. Appropriate volumes of homogenate serial dilutions were plated on LB agar plates to determine the number of CFUs. Mouse tissues were also collected and fixed in

4 % w / v

polyoxymethylene solution for

48 h

prior to histopathological examination via hematoxylin-eosin (HE) staining. Inflammation scores were assessed based on the number of pyogenic lesions in the hepatic lobules (liver), the number and size of the germinal centers (spleen), the integrity of the epithelial cell layer and inflammatory cell infiltration in the lamina propria (cecum). Mild inflammation was defined by scores ranging from 0 to 4, moderate inflammation was defined by scores ranging from 5 to 8, and severe inflammation was defined by scores ranging from 9 to 12.

Immunohistochemical analysis was performed with Histostain

​ T M

-SP Kits (Cat#SPN-9001; ZSbio, China) according to the manufacturer’s instructions. The cytokine levels in the cecum were evaluated using the enzyme linked immunosorbent assay (ELISA) detection kits of cytokine assay kits ELISA MAX

​ T M

Deluxe Mouse TNF-a (Cat#430904), IL-1β (Cat#432604), and IL-10 (Cat#431414; BioLegend, USA).

2.10. Statistical analysis

All the data were analyzed using GraphPad Prism 8.0 (GraphPad Software, USA). Student’s

t

test was used for comparing two groups of samples, while one-way analysis of variance (ANOVA) was used for analyzing data from more than three groups. Mouse survival data were evaluated using the log-rank test. The

P

values are indicated in the figures as follows: NS, not significant;

​ * * * P < 0.001

;

​ * * P < 0.01

; and

​ * P < 0.05

3. Results

3.1. LPD potently inhibits intracellular bacterial replication

To search for HDT drugs that are capable of eliminating intracellular bacteria, we performed a cell-based screening using ST as a model organism from an FDA-approved drug library consisting of 2320 compounds. After sufficient uptake of ST by RAW264.7 cells, the extracellular bacteria were killed by gentamycin, and intracellular bacterial growth in the presence of individual drugs was monitored for the indicated times (Fig. 1(a)). Among the screened drugs, we observed significantly decreased replication of ST SL1344 in cells treated with

40 μ g ⋅ m L - 1

gemcitabine

H C l

, ezetim-ibe, LPD, nitazoxanide, imidapril HCl, brexpiprazole, or indacaterol maleate (Table S4 in Appendix A). Among these positive hits, LPD exhibited the most significant inhibitory effect (Fig. 1(b)). A dose-dependent assay revealed that

4 μ g ⋅ m L - 1

LPD markedly reduced intracellular ST replication (Fig. 1(c)). A time-course assay also revealed a gradual decrease in the intracellular bacterial load in both macrophages and HeLa cells upon LPD treatment (Figs. 1(d) and (e)). Moreover, this reduction was further confirmed by immunostaining of the infected cells with an anti-Salmonella antibody (Figs. 1(f) and (g)). In addition to the influence of the ST, we further evaluated the influence of LPD on the growth of other bacterial species in RAW264.7 cells. Strikingly, the presence of

16 μ g ⋅ m L - 1

LPD in the infected samples significantly suppressed the replication of E. coli, S. flexneri, A. baumannii, L. monocytogenes, and S. aureus (Figs. 1(h)-(l)). Taken together, our data indicate that the FDA-approved LPD is an inhibitor of the intracellular growth of distinct intracellular bacterial pathogens.

3.2. LPD does not affect the in vitro growth or virulence of ST

Next, we assessed the direct antibacterial activities of LPD against the abovementioned bacterial species. The results showed that the MICs of LPD for ST, E. coli, S. flexneri, A. baumannii, L. mono-cytogenes, and

S

. aureus were greater than

1024 μ g ⋅ m L - 1

(Fig. S1(a) and Table S5 in Appendix A). Furthermore, in vitro growth curve analysis of ST showed that the presence of increasing concentrations of LPD had little effect on bacterial multiplication (Fig. S1(b) in Appendix A). Therefore, the observed inhibitory effect of LPD on the intracellular replication of bacteria was not attributed to its antibacterial effects.

The successful survival and proliferation of ST within host cells are strictly dependent on T3SS-1 and T3SS-2, which are encoded by Salmonella pathogenicity islands 1 and 2 (SPI-1 and SPI-2) [18,19], respectively. Both apparatuses deliver effector proteins into the host to assist in bacterial entry, prevent bacterial clearance by innate immunity, and establish an intracellular niche permissive for replication [18,19]. Therefore, it is likely that LPD suppresses T3SS-1 and/or T3SS-2, thus leading to the inhibition of intracellular replication of ST. To address this, we employed an established TEM reporter system to visualize the translocation of the T3SS-1 and T3SS-2 effectors [15,16]. Apparently, the translocation of SipA-TEM (a T3SS-1 substrate) or SopD2-TEM (a T3SS-2 substrate) into host cells was not affected by LPD treatment of ST, as indicated by the equal percentage of blue fluorescent cells detected in both samples (Figs. 2(a) and (b)). In line with this, the expression of SipA and the secretion of T3SS-1 effectors into the supernatant were not suppressed by the addition of various concentrations of LPD to the bacterial cultures (Figs. 2(c)-(e)). Taken together, our data suggest that the impairment of ST intracellular growth by LPD is not due to its effects on bacterial virulence.

3.3. Transcriptome regulation by LPD in ST-infected macrophages

These findings prompted us to focus on the alterations in host responses in cells challenged with intracellular bacteria after LPD treatment. To this end, we performed RNA sequencing and analyzed the DEGs among the different treatment groups. Infection of RAW264.7 cells with ST SL1344 led to the identification of 1089 DEGs compared to those in the blank control group (ST/ Blank), among which 515 genes were upregulated and 574 genes were downregulated (Fig. 3(a)). These changes may be critical for the pathogenesis of ST. In the ST-infected sample that received LPD (ST-LPD), 1369 differentially transcribed genes were observed in comparison with those in the ST sample, among which 738 genes were upregulated and 631 genes were downregulated (Fig. 3(a)). These alterations may represent the potential regulation of the host response by LPD to eliminate intracellular ST.

To evaluate the accuracy of the RNA sequencing data, ten DEGs were randomly selected for qRT-PCR validation. These included six genes (glutathione S-transferase, mu 1 (gstm1), sequestosome 1 (sqstm1), polo like kinase 3 (plk3), cathepsin D (ctsd), hematopoietic prostaglandin D synthase (hpgds), and alanyl aminopeptidase (anpep)) in the ST/Blank down group vs the ST-LPD/ST up group and four genes (ribonucleotide reductase M2 (rrm2), deoxycytidine triphosphate (dCTP) pyrophosphatase 1 (dctpp1), ribonucleotide reductase M1 (rrm1), and lymphotoxin B (ltb)) in the ST/Blank up group vs the ST-LPD/ST down group. Importantly, the trends in relative gene transcription in the blank, ST and ST-LPD groups determined by qRT-PCR were similar to those identified by RNA sequencing (Fig. S2 in Appendix A).

A total of 219 coexisting DEGs were identified by comparing all the groups (Fig. 3(a)): 123 in the ST/Blank down group vs the ST-LPD/ST up group and 96 in the ST/Blank up group vs the ST-LPD/ST down group. Interestingly, 8 out of the top 11 DEGs in the ST/Blank down group vs ST-LPD/ST up group were involved in the regulation of macrophage immunity, among which five genes (yippee like 3 (ypel3), cluster of differentiation (CD) 33 molecule (cd33), C-X-C motif chemokine receptor 4 (cxcr4), gpnmb, and gstm1) are regulators of lysosomal activity (Figs. 3(b) and (c)). Similarly, 8 out of the 12 top DEGs (septin2, regulator of G protein signaling 16 (rgs16), ltb, beta-1,4-glucuronyltransferase 1 (b4gat1), prostaglandin-endoperoxide synthase 2 (ptgs2), ISG15 ubiquitin like modifier (isg15), S100 calcium binding protein A8 (s100a8), interferon regulatory factor 7 (irf7), 2’-5’ oligoadenylate synthetase 1G (oas1g), histocompatibility 2, T region locus 24 (H2-T24), DEAD-box RNA helicase 58 (dhx58), and lectin, galactose binding, soluble 9 (lgals9)) in the ST/Blank up group vs ST-LPD/ST down group comparisons were involved in the regulation of macrophage immunity, among which seven genes are regulators of lysosomal activity (Figs. 3(d) and (e)). Taken together, these data raise the possibility that LPD triggers host immune responses to suppress the replication of invading bacteria.

3.4. LPD increases lysosomal activity in ST-infected macrophages

Lysosomes are acidic and hydrolytic organelles responsible for the digestion of macromolecules and are involved in innate immunity and tissue homeostasis through their ability to detect and eliminate microbes, debris, and dead cells in macrophages [20]. In addition, lysosomes function as sophisticated centers for signaling pathways involved in the response to nutrient and cellular stress [21]. The abovementioned results revealed the potential regulation of host immunity, especially lysosomal function, by LPD treatment. Indeed, further analysis of the RNA sequencing data showed that the transcription of 54 genes involved in the regulation of lysosomal activity and autophagy was increased in ST-infected macrophages that received LPD treatment (Fig. 4(a)). To validate the transcriptomic results, we employed LysoTracker and LysoSensor to probe the lysosomal function of macrophages. ST infection of host cells did not significantly alter lysosomal function (Figs. 4(b) and (c)). However, the fluorescence intensities of both LysoTracker and LysoSensor were substantially elevated in ST-infected cells treated with LPD (Figs. 4(b) and (c)). Therefore, it is likely that LPD-triggered lysosomal activity might contribute to the clearance of intracellular bacteria. Indeed, compared with infection in the control group, LPD treatment of ST-infected macro-phages yielded a markedly greater percentage of bacteria that acquired the lysosome marker LAMP-1 (Figs. 4(d) and (e)).

3.5. LPD promotes autophagy in ST-infected macrophages

Given the intimate connection between the autophagy process and lysosomes as well as the effect of LPD on lysosomal acidification, we suspect that LPD treatment may also regulate the host autophagy response. To test this hypothesis, we detected the levels of microtubule-associated protein 1 light chain 3 alpha (LC3) and p62 in RAW264.7 cells by western blot analysis using specific antibodies. Remarkably, while the control cells or ST-infected cells only presented basal levels of LC3-II, the lipidated form of LC3, and the addition of LPD to the cell culture increased LC3-II levels to levels comparable to those of rapamycin-treated cells (Figs. 5(a) and (b)). The ubiquitin-binding scaffold protein p62 is degraded upon autophagy induction and is often used as a marker of autophagic flux. Surprisingly, the amount of p62 in cells receiving LPD was slightly upregulated compared to that in the untreated control cells (Figs. 5(a) and (c)). This inconsistency might be attributable to the LPD-induced increase in the transcription of p62 (Fig. S2). In addition, the induction of autophagy by LPD was also observed in HeLa (Figs. 5(d)-(f)), MAC-T (Figs. 5(g)-(i)), and Caco-2 cells (Figs. 5(j)-(l)).

KEGG analysis of DEGs between the ST/Blank up and ST-LPD/ST comparisons revealed that nine genes were involved in the phos-phatidylinositol 3-kinase (PI3K)/AKT serine/threonine kinase (Akt) signaling pathway (Fig. S3 in Appendix A). The mTOR is a downstream target of the PI3K/Akt signaling pathway [22] and plays a crucial role in regulating the autophagy process [23]. In view of these findings, we further evaluated the phosphorylation status of mTOR (p-mTOR) in ST-infected RAW264.7 cells upon treatment with LPD. Strikingly, cells challenged with ST exhibited an increase in p-mTOR, while LPD treatment decreased the p-mTOR levels of the ST infected samples (Figs. 5(m)-(p)). Therefore, LPD-induced autophagy in ST-infected macrophages likely occurs via inhibition of the PI3K/Akt/mTOR pathway.

Next, we investigated whether LPD-stimulated autophagy contributed to the clearance of intracellular ST. To achieve this goal, we first infected RAW264.7 cells with ST and subsequently treated them with LPD. After immunostaining the infected samples with anti-Salmonella and anti-LC3 antibodies, we observed a significantly greater percentage of bacteria that were surrounded by LC3 puncta (Figs. 6(a) and (b)). Furthermore, the addition of 3-methyladenine (3MA), an autophagy inhibitor, to the infected sample relieved the LPD-mediated inhibition of ST intracellular replication (Fig. 6(c)). Taken together, our data indicate that LPD inhibits the intracellular proliferation of ST by promoting autophagy.

3.6. GPNMB is critical for LPD-induced autophagy and clearance of intracellular ST

According to the RNA sequencing data, gpnmb had the highest transcription among the DEGs. gpnmb expression markedly decreased upon ST infection but increased after LPD treatment (Fig. 7(a)). The gpnmb gene encodes GPNMB, a highly glycosylated protein localized either on the cell membrane or in endosomes or lysosomes [24]. GPNMB is involved in the regulation of various host cell physiological processes, such as cell migration, proliferation, differentiation, and adhesion [25]. Furthermore, GPNMB was reported to regulate host autophagy and lysosomal function[26,27].

To further dissect the mechanism of the LPD-mediated clearance of intracellular bacteria, we first validated the transcriptional levels of gpnmb among the experimental groups by qRT-PCR (Fig. 7(b)). Infection of RAW264.7 cells with ST, S. flexneri or E. coli reduced gpnmb transcription, while treatment of the infected cells with LPD significantly enhanced the transcriptional levels of gpnmb (Fig. 7(c)). Moreover, western blot analysis of macrophage lysates using specific antibodies also revealed an increase in the amount of GPNMB upon LPD treatment (Figs. 7(d) and (e)). Taken together, these data suggest that GPNMB may be crucial for the elimination of various intracellular pathogens by LPD.

Next, we employed RNA interference (RNAi) technology to knock down GPNMB expression in RAW264.7 cells (Figs. 8(a) and (b)). Importantly, although LC3-II and p62 levels were not affected by GPNMB knockdown, the increase in LC3-II and p62 expression triggered by LPD was significantly attenuated by the lack of GPNMB (Figs. 8(c)-(e)). Moreover, we found that knockdown of gpnmb had no impact on the intracellular growth of ST. However, the inhibition of intracellular ST replication induced by LPD was markedly alleviated by siRNA-gpnmb (Fig. 8(f)). Therefore, these data indicate that GPNMB plays a vital role in the LPD-mediated promotion of autophagy and clearance of intracellular bacteria.

3.7. LPD effectively protects against ST infection in G. mellonella and mouse models

The inhibitory effect of LPD on intracellular bacterial replication has illustrated its therapeutic potential for bacterial infections. To address this, we established G. mellonella and mouse ST infection models. In the G. mellonella model, all the larvae died within

12 h

after inoculation with

1 × 105

CFUs of ST. Strikingly, the administration of larvae with a single dose of LPD dissolved in

10 μ L

150 μ g ⋅ k g - 1

improved the survival rate to

40 %

(Fig. 9(a)). To measure the survival rate of the mice in the infection model, each mouse was orally inoculated with

1 × 107

CFUs of ST after the administration of streptomycin in the drinking water for three days. Then, the mice were treated with

5 m g ⋅ k g - 1

LPD at

12 h

intervals until the endpoint of the study. All mice in the infection group that were administered PBS died on the ninth day, while mice in the LPD treatment group showed 50% survival (Fig. 9(b)).

To assess the bacterial burden in the organs, necropsy, and histopathology analysis, each mouse was orally inoculated with

5 × 106

CFUs of ST and treated with LPD on the same schedule as described for the survival rate test. Mice were sacrificed on the fifth day post infection. The necropsy results showed that the cecum contained less solidified feces in the ST infection group, whereas healthy and solidified feces were observed in the LPD-treated group (Fig. 9(c)). In addition, the administration of LPD to mice resulted in significantly decreased bacterial colonization in both the liver and spleen (Fig. 9(d)). Finally, the histopathological changes were monitored via HE staining of the organs. The epithelial cell layer of the cecum was severely exfoliated in ST-infected mice, while LPD treatment led to less epithelial shedding. ST causes extensive inflammatory cell infiltration, congestion, and bleeding in the livers of mice. However, LPD administration significantly alleviated these pathological changes (Fig. 9(e)). Additionally, the spleens of the ST-infected mice that received LPD treatment exhibited markedly less inflammatory cell infiltration than those of the infection control mice (Fig. 9(e)). The inflammatory scores were calculated by summing the scores of the liver, spleen, and cecum. The results demonstrated a significantly lower level of inflammation in the LPD treatment group than in the ST infection group (Fig. 9(f)). Taken together, our data indicate that LPD could effectively protect against ST infection in animal models.

In addition, phosphorylated mTOR and p62 in the mouse cecum were analyzed via immunohistochemistry. Cecal p-mTOR expression in LPD-treated mice was significantly lower than that in ST-infected mice (Figs. S4(a) and (b) in Appendix A), while the expression of p62 in LPD-treated mice was significantly greater than that in ST-infected mice (Figs. S4(c) and (d) in Appendix A). Therefore, LPD treatment promoted autophagy in ST-infected mice, which is consistent with the observations observed in cell lines.

3.8. LPD alleviates the intestinal inflammation caused by ST infection

The RNA sequencing data revealed 34 DEGs involved in inflammation regulation, including cytokines, chemokines, chemokine receptors, and Toll-like receptors (Fig. 10(a)). These data suggested that, in addition to clearing intracellular bacteria by promoting the host autophagy process, LPD might modulate the host inflammatory response to facilitate elimination of invading bacteria. To this end, we first measured the transcriptional levels of cytokines in RAW264.7 cells by qRT-PCR. Following ST infection, the messenger RNA (mRNA) levels of tumor necrosis factor (TNF)-

α

, interleukin (IL)-6, IL-1

β

, and interferon (IFN)-

γ

were strikingly upregulated (Figs. 10(b)-(e)). Importantly, the addition of LPD to the infected cells caused a marked reduction in the transcript levels of TNF-

α

, IL-6, and IL-1

β

(Figs. 10(b)-(d)). However, the transcription of IFN-

γ

was not affected by LPD (Fig. 10(e)). In contrast, ST infection suppressed IL-10 transcription, and LPD treatment significantly restored the mRNA levels of IL-10 (Fig. 10(f)). Furthermore, we collected cecal samples from ST-infected mice and measured cytokine levels via ELISA. ST infection elevated TNF-

α

and IL-1

β

levels (Figs. 10(g) and (h)) but reduced the amount of IL-10 (Fig. 10(i)). Importantly, LPD treatment of ST-infected mice significantly reduced (TNF-

α

and IL-1

β

) (Figs. 10(g) and (h)) or increased (IL-10) (Fig. 10(i)) cytokine levels to those of uninfected mice. Collectively, these data indicate that LPD inhibits proinflammatory cytokines and elevates anti-inflammatory cytokines to promote therapeutic outcomes.

4. Discussion

Intracellular bacterial pathogens such as

S

. Typhimurium,

M

. tuberculosis, and S. flexneri can invade and reside in host cells, including epithelial cells and macrophages, thus protecting themselves from killing by both the host immune system and antibiotics. These intracellular bacteria can cause acute life-threatening infections as well as recurring chronic infections. Canonical antimicrobial therapeutics for treating intracellular bacterial infection are challenging due to the low permeability and rapid intracellular degradation of most antibiotics, which greatly decrease antimicrobial efficacy [28,29]. Increasing the dosage might improve the therapeutic effects to a certain degree, but it also enhances economic burdens and magnifies off-target impacts on the normal flora [30]. Moreover, this dilemma has been strongly aggravated by the emergence and dissemination of multidrug-resistant bacterial strains. Hence, the development of novel anti-infection drugs or strategies is urgently needed.

Although only 12 new antibiotics have been approved for use in the 21st century, identifying novel and effective antibiotics or improving the treatment efficacy of existing antimicrobials by pharmaceutical means are the first choices for controlling bacterial infections [31]. With advances in new technologies, such as culturo-mics and genomics [32], several promising antimicrobial compounds (e.g., teixobactin [33] and darobactin [34]) have been identified in recent years. Despite this progress, additional approaches are needed to supplement antimicrobial therapeutics, and the HDT strategy, which aims to boost host antimicrobial mechanisms, has attracted increased interest. Unlike traditional antibiotics, HDT drugs do not possess direct bactericidal or bacteriostatic activities, thus imposing less selective pressure on bacteria [10]. Therefore, HDT drugs are less likely to cause resistance and can maintain the integrity of the microbiome [30]. To successfully parasitize host cells, intracellular bacteria not only exploit cellular nutrition for replication but also need to compromise host defensive mechanisms that are detrimental to bacterial survival. Consequently, the HDT strategy is particularly suitable for the eradication of intracellular pathogens. Importantly, advances in the understanding of host-pathogen interactions have provided directions for the design or screening of HDT drugs [35]. In particular, the development and application of CRISPR/Cas genome editing has allowed for large-scale screening of host factors and processes required for intracellular bacterial survival and replication [36,37]. Currently, clearance of intracellular bacteria by HDT can be achieved by blocking host pathways essential for bacterial survival or activating pathogen-antagonizing mechanisms.

Among the host processes, autophagy not only plays important roles in maintaining cellular homeostasis by providing energy sources but also acts as a critical innate defense mechanism termed xenophagy [38] to eliminate invading microbes. However, many intracellular pathogens can avoid xenophagic clearance via specific virulence determinants. For example, ST utilizes the T3SS- 2 effector protein SopF to inactivate the xenophagic response via ADP-mediated ribosylation of essential factors involved in the signaling pathway [11]. Therefore, reinstalling or activating autophagy signaling represents a promising and important method for HDT drug development for treating intracellular infections [13]. Indeed, previous studies have revealed multiple chemical compounds that can protect host cells from intracellular bacterial infection, especially

M

. tuberculosis infection, by modulating host autophagy [39]. For instance, the epidermal growth factor inhibitor gefitinib acts as an autophagy inducer to inhibit

M

. tuberculosis replication [40]; metformin, the first-line drug for type 2 diabetes treatment, is also effective at eliminating intracellular

M

. tuberculosis by triggering autophagy [41]. Considering its acceptable toxicity and broad application, metformin has served as a promising anti-TB drug that is ready for clinical tests [42]. Additionally, Chiu et al. [43] demonstrated that a novel small molecule agent, AR-12, eradicates intracellular ST by promoting autophagosome formation in infected macrophages. Administration of AR-12 to ST-infected mice could significantly reduce hepatic and splenic bacterial colonization and prolong survival, illustrating its potential application in controlling Salmonella infection [43].

In this study, through a cell-based screening of FDA-approved drugs, we demonstrated that LPD is a potent inhibitor of intracellular ST. Treatment of host cells with LPD could markedly increase the host autophagy response as well as lysosomal activity, thereby mediating host eradication of intracellular bacteria. Importantly, LPD showed promising therapeutic potential against ST infection, as proven by the increased survival rate, reduced bacterial load in organs, and alleviated histopathological damage in animal infection models. Since blocking the host autophagic response is one of the common mechanisms employed by various bacteria to survive and replicate within host cells [44], LPD treatment could also inhibit the intracellular replication of E. coli, S. flexneri, A. baumannii, L. monocytogenes, and Staphylococcus aureus. Moreover, LPD was also shown to inhibit the replication of Middle East respiratory syndrome coronavirus (MERS-CoV), severe acute respiratory syndrome (SARS) coronavirus, and human coronavirus 229E in cell culture [45]. Therefore, our study suggested that LPD is an HDT candidate drug that possesses broad-spectrum activity against distinct intracellular pathogens. In addition, previous studies have demonstrated that LPD can augment the activity of the antibiotics minocycline [46] and colistin [47], indicating the potential application of LPD as an adjunct to canonical chemotherapy for treating intracellular infections. LPD is a nonprescription

μ

-opioid receptor agonist that is widely used as an antidiarrhea agent in clinical practice [48]. Since LPD has a low penetration rate in the central nervous system (CNS), it is generally considered safe at therapeutic doses [49]. However, high-dose LPD can potentially cause cardiac toxicity [50] and affect the CNS [51]. In addition, other adverse effects, such as constipation and abdominal pain, can occur occasionally following the administration of LPD [49]. Therefore, the safety implications of LPD need to be re-evaluated before repurposing LPD for the treatment of other diseases.

GPNMB is a transmembrane glycoprotein that participates in the regulation of multiple host processes [25]. For instance, GPNMB enhances autophagy via suppression of mTOR signaling [27]. In addition, GPNMB has become a biomarker for lysosomal dysfunction in macrophages [24]. Interestingly, one recent study revealed that GPNMB restricts respiratory syndrome virus (PRRSV) replication by inhibiting the fusion of autophagosomes and lysosomes [52]. Our mechanistic study showed that the effect of LPD on the clearance of intracellular bacteria is dependent on the LPD-induced increase in the expression of GPNMB. Taken together, these findings indicate that GPNMB might be an ideal drug target for HDT aimed at promoting host autophagy.

Acknowledgment

This work was supported by the National Key Research and Development Program of China (2021YFD1801000), the Natural Science Foundation of China (32373066), the Natural Science Foundation of Jilin Province (20230101142JC), and the Fundamental Research Funds for the Central Universities.

Compliance with ethics guidelines

Hongtao Liu, Siqi Li, Le Deng, Zhenxu Shi, Chenxiao Jiang, Jingyan Shu, Yuan Liu, Xuming Deng, Jianfeng Wang, Zhimin Guo, and Jiazhang Qiu declare that they have no conflict of interest or financial conflicts to disclose.

Data availability

The RNA sequencing data used to support our study findings are original and stored in the Gene Expression Omnibus (GEO) of the National Center for Biotechnology Information (NCBI)

​ †

under accession number GSE236340. The following secure token was created to allow review of the GSE236340 dataset while remaining private: itwtqaqenfsjzil.

Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.eng.2024.01.011.

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